Intercellular nanotubes: insights from imaging studies and beyond

Abstract

Cell-cell communication is critical to the development, maintenance, and function of multicellular organisms. Classical mechanisms for intercellular communication include secretion of molecules into the extracellular space and transport of small molecules through gap junctions. Recent reports suggest that cells also can communicate over long distances via a network of transient intercellular nanotubes. Such nanotubes have been shown to mediate intercellular transfer of organelles as well as membrane components and cytoplasmic molecules. Moreover, intercellular nanotubes have been observed in vivo and have been shown to enhance the transmission of pathogens such as human immunodeficiency virus (HIV)-1 and prions in vitro. These studies indicate that intercellular nanotubes may play a role both in normal physiology and in disease.

Figure 1

Intercellular nanotubes. (1) Growing cellular nanotubes (cytonemes) in culture of fragments cut from Drosophila wing discs. Fluorescence microscopy initially revealed round green fluorescent protein (GFP)-containing cells, (a), but after approximately 40 min in culture, multiple small processes containing GFP emerged (green). (b), These processes extended and retracted rapidly. After approximately 60 min, the cells extended long, GFP-containing processes, (c, d). The cytonemes were found to grow unidirectionally toward C-fragment cells (nonfluorescent) (d, arrowheads). Scale bar, 10 μm. (Reprinted with permission from Ref Ref 2. Copyright 1999 Elsevier). (2) The architecture of tunneling nanotubes (TNTs) between cultured rat pheochromocytoma (PC12) cells was analyzed by 3D live-cell microscopy. Cells were connected with surrounding cells via one, (a), or several TNTs, (b). Rarely, branched TNTs were observed (c, arrow). A selected (x–z) section obtained from a confocal 3D reconstruction illustrates how TNTs do not adhere to the substrate, (d), scale bar, 15 μm. (e), The ultrastructure of TNTs in PC12 cells was analyzed by scanning electron microscopy, scale bar, 10 μm, E1-F3: 200 nm. (Reprinted with permission from Ref 4. Copyright 2004 AAAS). (3) Membrane nanotubes connect immune cells. A nanotube connecting Epstein–Barr virus-transformed human B-cells (a) where the fluorescence image (right) shows emission from glycosylphosphatidylinositol anchored GFP (green). (b) Membrane nanotube between two primary human macrophages. Scale bar, 10 μm. (Reprinted with permission from Ref 5. Copyright 2004 AAI). (4) An artificial cellular nanotube with schematic representation showing the formation of a daughter vesicle from a cell membrane bleb. Membrane blebs were induced in NG108-15 cells by a combination of DTT and formaldehyde; subsequently, one bleb was electroinjected with a buffer-filled micropipette. Following translation of the micropipette, a nanotube connection was formed and the injected buffer led to the formation of a daughter vesicle at the micropipette tip. Organelles and cytoskeletal structures remained within the cell, while the bleb most likely enclosed low molecular weight cytosolic molecules. Both the bleb and the daughter vesicle have membrane proteins embedded in the membrane. (Adapted with permission from Ref 6. Copyright 2007 ACS).

Figure 2

Pathogens exploit intercellular nanotubes (ICNs). (1) Viral particles. ICNs mediate cell-to-cell transmission of retroviruses. (a), Superimposed video frames illustrate the opposing movements of a receptor-expressing filopodium (red) and the movement of viral particles (green). Scale bar, 5 μm. (b), Time-lapse sequence of viral particles moving from the infected cell toward the noninfected target cells. The moving viral particle (green) was colocalized with mCAT1 receptor (red). (c), The viruses move along the outer surface of filopodial bridges toward target cells and correlated to single approximately 100-nm particles observed by SEM (black arrows). Scale bar, 500 nm. (a-c, Adapted with permission from Ref 16. Copyright 2007 Macmillan Publishers Ltd.). (d), Cellular nanotubes can be induced by human immunodeficiency virus (HIV)-infection of macrophages. Distinct HIV vesicles localized in nanotubular processes: actin (red), HIV-p24 (green), and DAPI (blue). Arrows denote HIV-p24 positive vesicles being transported across long nanotubes. (Adapted with permission from Ref 15. Copyright 2009 Elsevier). (2) Bacteria. Bacteria can surf along membrane nanotubes, aided by constitutive flow of nanotube surface. (a), Brightfield time-lapse sequence showing Mycobacterium bovisBCG expressing soluble green fluorescent protein (GFP) incubated with human macrophages. A cluster of bacteria (arrowhead) is shown trapped on an ICN connecting two macrophages and transported along the nanotube to the cell body where they were phagocytosed. (b), To confirm that the bacteria were indeed internalized, the cell membrane was labeled by the addition of membrane dye directly to the imaging chamber, schematic view to the left. (Adapted with permission from Ref 11. Copyright 2004 AAI). (c), Schematic illustrating the principle of extratubular Marangoni transport of bacteria over nanotubes in nanotube–vesicle networks (NVNs). Bacteria attached to the outer leaflet of the vesicle bilayer were transported over the nanotube by membrane flow. (d–f), Micrograph sequence demonstrating the surfing of bacteria between neighboring vesicles in NVNs. Scale bar, 20 μm. (Reproduced with permission from Ref 36. Copyright 2008 RSC). (3) Prions. Prions transfer through ICNs. (a), A three-dimensional reconstruction showing an ICN (yellow arrow) connecting a Cath.a-differentiated central nervous system cell (CAD cell) transfected to express actin-GFP (green) and labeled with LysoTracker (red) and an untransfected CAD cell. (b), A video sequence captures a vesicle moving inside a nanotube [termed tunneling nanotube (TNT)] and entering the cytoplasm of the recipient cell. (c), Diseased prions (PrPSc) were found in vesicular structures (white arrows) inside TNTs, as well as in the cytoplasm of the transfected cell, showing that PrPSc can transfer through TNTs. (d), Quantification of endogenous PrPSc transfer from prion infected CAD cells (ScCADs) to CAD cells through TNTs. CAD cells (red) cocultured with ScCAD (1–3) or with ScCAD in the presence of latrunculin to block TNT formation (4). Efficient transfer of PrPSc is detected only in cells connected through TNTs: (1), CAD cells not touching ScCAD cells; (2), CAD cells in direct cell contact with ScCAD cells; (3), CAD cells in contact with ScCAD cells through TNTs; (4), latrunculin-treated cocultures. Scale bar, 10 μm. (Adapted with permission from Ref 18. Copyright 2009 Macmillan Publishers Ltd.).

Figure 3. In vivo

intercellular nanotubes (ICNs) observed between cells in the corneal stroma tissue of mice. (a), Chimeric mouse corneal whole mount reveals a donor-derived (green fluorescent protein (GFP)⁺ /green) major histocompatibility complex (MHC) class II⁺ (red) and double positive (yellow) cell connected via a fine ICN (arrows) to a resident MHC class II⁺ GFP^- cell (red only). (b), Two donor-derived MHC class II⁺ cells expressing varying amounts of GFP joined by a fine, straight ICN. (c) and (d), Long, nonbridging membrane nanotubes on MHC class II+ cells in the naive, (c), and inflamed, (d), mouse corneal stroma. Scale bar, 20 μm; inset scale bar, 10 μm. (Reprinted with permission from Ref 19. Copyright 2008 AAI).

Figure 4

Cytoskeletal structure and formation of ICNs (1) Actin structure of ICNs illustrated with macrophages, (a), differential interference contrast micrograph, (b), micrograph showing a 3D rendered confocal stack. F-actin stained by phalloidin (white) and the nucleus stained with Hoescht (red). (c), A zoomed-in region showing the phalloidin stained nanotubes. (2) Formation of ICNs. (a–c), Bridging nanotubes. (a), Phase contrast image of live T24 cells, (b), fluorescence micrograph with actin labeling of the same cells as in A. The white arrows in (a) and (b) indicate short and dynamic membrane protrusions with which the approaching cell explores its surroundings. The black arrow in, A, points at protrusions that have already connected to the target cell. Bridging nanotubes can be more than 20 μm in length and bifurcations are occasionally observed (arrow in C). (Reprinted with permission from Ref 51. Copyright 2008 Elsevier). (d), Time-lapse imaging of 721.221 cells forming a transient contact demonstrates that a connecting nanotube can form as cells move apart after contact. Scale bar, 10 μm. (Reprinted with permission from Ref 5. Copyright 2004 AAI). (3) Schematic illustration, (a), of stabilization of nanotubular membrane protrusions by accumulation of anisotropic membrane nanodomains in the tubular region. The growing actin filaments push the membrane outward. The protrusion could additionally be stabilized by accumulated anisotropic nanodomains that favor an anisotropic cylindrical geometry of the membrane. The cylindrical-shaped anisotropic membrane domains, once assembled in the membrane region of a nanotubular membrane protrusion, keep the protrusion mechanically stable even if the cytoskeletal components (actin filaments) are disintegrated. (Reprinted with permission from Ref 51. Copyright 2008 Elsevier). (b), The general regulatory pathway of actin polymerization in filopodia (4) LFA-1 activation is responsible for two distinct pathways of nanotube formation in Jurkat T-cells. (a), Example of the extension and nanotube outgrowth by Cathepsin X-up regulated Jurkat T-cells in a three-dimensional environment, leading to increased ICN-mediated cell-to-cell contact. (b), Wt cells remained in a spherical shape. Nanotubes form upon uropod elongation in the absence of prior intercellular contact. (c), The proposed mechanism of nanotube outgrowth via persistent LFA-1 activation (mediated by cathepsin X). Talin binds to the cytoplasmic tail of LFA-1, followed by binding of ICAM-1 to the extracellular domain, enabling inside-out driven actin reorganization, uropod formation, elongation, and ICN formation. (d), The proposed mechanism of nanotube formation following intercellular contact between T-cells. LFA-1/ICAM-1 interactions arising in T-cell aggregation are enhanced in cathepsin X-upregulated Jurkat cells. Prolonged LFA-1 activation enables cytoskeletal reorganization, similar to that associated with uropod outgrowth, with talin-binding to the cytoplasmic tail of LFA-1 and subsequent membrane nanotube formation as cells depart. (Modified with permission from Ref 52. Copyright 2009 Springer Science+Business Media).

Figure 5

Membrane continuity of intercellular nanotubes. (a) Transmission electron microscope (TEM) ultrastructure of a tunneling nanotube (TNT) between rat pheochromocytoma cells where a serial sectioning showed that, at any given point along TNTs, the membrane appeared to be continuous. From Rustom et al. reprinted with permission from AAAS. (b) TEM ultrastructure of an ICN between T-cells reveals a closed border i.e., no direct cytoplasmic contact or membrane mixing. (b′, b″) Insets show higher magnifications of the nanotube and the junction. Scale bar, 500 nm. (Reprinted with permission from Ref 17. Copyright 2008 Nature Publishing Group).

Figure 6

Schematic illustration of the nanotube-vesicle network fabrication principle. (a), By a combination of mechanical deformation and electric pulses across the liposome, a microinjection needle is inserted into a unilamellar liposome connected to a multilamellar protrusion (not shown). (b), After the lipid has adhered to the injection needle, the micropipette is pulled away from the liposome. (c), A lipid nanotube is created between the tip of the micropipette and the liposome. (d), By applying a low pressure in the microinjection needle, ejected liquid expands the nanotube into a liposome at the microinjection tip, transferring additional lipid material from the multilamellar liposome (not shown). (e), After the liposome has reached a desired size, it is allowed to adhere to the surface. Thereafter, the micropipette is removed by applying electric pulses and simultaneously pulling it out of the liposome.

Figure 7

Membrane dynamics and three-way junction formation. (1) Direct nanotube extraction from cell by laser trap manipulation. (a), As a membrane nanotube is extracted from the cell, force versus microscope stage displacement, (b), is recorded, where the dashed line indicates the plateau average of the force. (2) Transition from a V to a Y shape of cellular nanotubes. (a), Before bifurcation, (b), after bifurcation, and, (c), the corresponding force versus microscope stage displacement. Points corresponding to images (a) and (b) are marked by arrows; the bifurcation position coincides with the dip in the curve seen in image (c). (Reprinted with permission from Ref 50. Copyright 2008 Springer Science+Business Media). (3) Model systems such as nanotube–vesicle networks can provide insight into membrane properties. (a–c), Inverted fluorescence images showing merging of nanotubes. Based on observations of the surfactant flow (black arrows, 1–3) on the nanotubes in model membrane systems, it is shown that a Y-junction propagates with a zipper-like mechanism. The surfactants from two nanotube branches undergo 1:1 mixing at the junction, and spontaneously form the extension of the third nanotube branch. Scale bar, 30 μm. (d), Experimental data for the total nanotube length of four different Y junctions, and fits obtained by numerical integration of the fluid-string model. The value in the inset is the nanotube string tension estimated from the fits. (Adapted with permission from Ref 57. Copyright 2006 American Physical Society).

Figure 8

Materials and methods of nanotubular studies. (1) Fluorescent imaging techniques such as confocal imaging provide spatiotemporal information based on fluorescent staining. Labeling of cells is based on, (a), genetically modified cell lines expressing fluorescent proteins, (b), membrane stains, or, (c), antibodies. The panels to the right give specific examples of nanotubes tagged using these methods. (d), B-cells expressing glycosylphosphatidylinositol anchored GFP. (e), nanotube-vesicle network stained with the membrane dye DiO. (f), Primary macrophages stained with phalloidin for F-actin (red) and with monoclonal antibody against α-tubulin (green). (2) Electron microscopy provides structural details on a nanometer resolution scale. (a), Scanning electron microscopy can provide external structural information. Adapted from Veranic et al. © 2008, with permission from Elsevier. (b), Transmission electron microscopy illuminates the internal structure, such as membrane junctions. (Adapted with permission from Ref 51. Copyright 2008 Nature Publishing Group). (3) Femtoliter- and picoliter-sized droplets as nanolaboratories for manipulating single cells and subcellular compartments is a potential route to analyze cellular nanotubes. A potential route to analyze intercellular nanotubes could be the use of droplet-nanolaboratories as has been demonstrated for single cells and subcellular compartments. (Reprinted with permission from Ref 81. Copyright 2009 ACS).

Figure 9

Transport of cargo in intercellular nanotubes (ICNs). (1) Diffusional translocation of molecules is effective on the cellular scale. (a) and (b), Time sequence of the spread of calcium signal (pseudocolored) on the left and the corresponding differential interference contrast image on the right. (Reprinted with permission from Ref 8. Copyright 2005 Elsevier). (c–g), Model systems for studies of diffusional effects in a nanotube–vesicle network (NVN). Diffusive transport rates were controlled with a thermoactuated hydrogel valve (in container 2). (Adapted with permission from Ref 95. Copyright 2008 ACS). (2) Constitutive membrane flow and Marangoni transport are tightly linked to actin polymerization and membrane tension. (a), Vesicular gondolas formed as an integral part of ICNs in the human urothelial cell line. (b), Fusion of a gondola (arrows) with a cell body, showing directional movement of the gondola along a nanotube. (Adapted with permission from Ref 51. Copyright 2008 Elsevier). Gondola formation in NVN systems. (c), Schematic for the formation of nanotube-integrated mobile vesicles in NVNs. (d), A difference of membrane tension induces Marangoni transport between vesicles: by pressing a vesicle with a microfiber, a flow toward the tense vesicle is created (σ2 > σ1). (e), Stationary state of a lipid tube connecting tense and floppy vesicles. (f), By selective manipulation of a node in NVNs, introduced gondolas can be delivered to a vesicle of choice. (g), Time-sequences of the formation of nanotube-integrated mobile vesicles. (3) Transportation along and elongation of nanotubes are regulated by actin polymerization and molecular motors. (a–d), Virus cell surfing is actin and myosin II dependent. (Reprinted with permission from Ref 35. Copyright 2005 originally published in JCB). (e–g), An example where microtubule connected membrane tubes are formed from giant vesicles by dynamic association of motor proteins (Reprinted with permission from Ref 96. Copyright 2006 National Academy of Sciences).